Correction

NEUROSCIENCECorrection for “Single-nucleus RNA sequencing of mouse au-ditory cortex reveals critical period triggers and brakes,” by Brian T.Kalish, Tania R. Barkat, Erin E. Diel, Elizabeth J. Zhang,Michael E. Greenberg, and Takao K. Hensch, which was firstpublished May 13, 2020; 10.1073/pnas.1920433117 (Proc. Natl.Acad. Sci. U.S.A. 117, 11744–11752).The authors note that the following statement should be in-

cluded at the end of the legend for Fig. 3: “Some of the data usedin B, C, and D are readapted from the C57 dataset used in ref.9.” The authors also note that the following statement should beincluded at the end of theMaterials and Methods section, on page11751: “C57 data were readapted from ref. 9 in accordance withthe Institutional Animal Care and Use Committee guidelines toreduce unnecessary use of animals. Statistics were taken internalto each genotype.” The authors apologize for the oversight in notclearly identifying the dataset from their prior publication.

Published under the PNAS license.

First published August 3, 2020.

www.pnas.org/cgi/doi/10.1073/pnas.2014145117

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https://www.pnas.org/site/aboutpnas/licenses.xhtmlhttps://www.pnas.org/cgi/doi/10.1073/pnas.2014145117https://www.pnas.org

Single-nucleus RNA sequencing of mouse auditorycortex reveals critical period triggers and brakesBrian T. Kalisha,b,1, Tania R. Barkatc,d,1,2, Erin E. Dielc,d,1, Elizabeth J. Zhangd, Michael E. Greenberga,3,and Takao K. Henschc,d,e,f,3

aDepartment of Neurobiology, Harvard Medical School, Boston, MA 02115; bDivision of Newborn Medicine, Department of Medicine, Boston Children’sHospital, Boston, MA 02115; cDepartment of Molecular and Cellular Biology, Harvard University, Cambridge, MA 02138; dCenter for Brain Science, HarvardUniversity, Cambridge, MA 02138; eF. M. Kirby Neurobiology Center, Department of Neurology, Boston Children’s Hospital, Harvard Medical School, Boston,MA 02115; and fChild Brain Development, Canadian Institute for Advanced Research, Toronto, ON M5G 1M1, Canada

Contributed by Michael E. Greenberg, March 2, 2020 (sent for review November 27, 2019; reviewed by David F. Clayton and Étienne de Villers-Sidani)

Auditory experience drives neural circuit refinement during win-dows of heightened brain plasticity, but little is known about thegenetic regulation of this developmental process. The primaryauditory cortex (A1) of mice exhibits a critical period for thalamo-cortical connectivity between postnatal days P12 and P15, duringwhich tone exposure alters the tonotopic topography of A1. Wehypothesized that a coordinated, multicellular transcriptionalprogram governs this window for patterning of the auditorycortex. To generate a robust multicellular map of gene expression,we performed droplet-based, single-nucleus RNA sequencing(snRNA-seq) of A1 across three developmental time points (P10,P15, and P20) spanning the tonotopic critical period. We also tone-reared mice (7 kHz pips) during the 3-d critical period and collectedA1 at P15 and P20. We identified and profiled both neuronal (glu-tamatergic and GABAergic) and nonneuronal (oligodendrocytes,microglia, astrocytes, and endothelial) cell types. By comparingnormal- and tone-reared mice, we found hundreds of genes acrosscell types showing altered expression as a result of sensory ma-nipulation during the critical period. Functional voltage-sensitivedye imaging confirmed GABA circuit function determines criticalperiod onset, while Nogo receptor signaling is required for itsclosure. We further uncovered previously unknown effects of de-velopmental tone exposure on trajectories of gene expression ininterneurons, as well as candidate genes that might execute tono-topic plasticity. Our single-nucleus transcriptomic resource of de-veloping auditory cortex is thus a powerful discovery platformwith which to identify mediators of tonotopic plasticity.

auditory cortex | GAD65 | Nogo receptor | single-cell sequencing

Activity-dependent plasticity shapes neural circuits in re-sponse to sensory experience during distinct developmentalwindows, termed “critical periods.” Heightened plasticity atthese times refines the anatomic and functional architectureacross brain regions (1). Decades of work in the primary visualcortex (V1) have defined how activity drives molecular and cel-lular events to open and close critical periods (2–4). Far lessdetail is known in other sensory systems, such as the primaryauditory cortex (A1), where biased early-life acoustic exposureslead to robust alterations in the spatial organization of a topo-graphic map of sound frequency, called tonotopy (5). Exposureto particular tones (tone-rearing) or language-specific pho-nemes early in life can shift auditory tuning curves and per-ception in favor of the experienced acoustic environment atthe expense of spectrally neighboring frequencies or speechsound contrasts (6, 7).Molecular mechanisms underlying critical period plasticity

have been most extensively studied for visual acuity (8). Incontrast, tonotopic plasticity occurs earlier and for a shorterperiod of time as compared to the V1 critical period for oculardominance (9–11). Nevertheless, both critical periods shareseveral features in common. Both auditory and visual corticalplasticity are accompanied by morphological remodeling of

dendritic spines (9, 12). Plasticity in A1 emerges earlier than inV1 in close register with accelerated maturation of parvalbumin-positive (PV) cells, shown to be pivotal for critical period onsetin V1 (4, 13). Closure of the auditory critical period can bedelayed or reopened in adulthood by exposure to continuousbroadband noise (0.8 to 30 kHz), which induces several changesin A1 associated with a more plastic state similar to that found indark-reared V1 (14, 15). Specifically, extended A1 plasticity isassociated with decreased PV and BDNF expression, as well asfewer GABAA α1 and β2/3 subunits (16, 17).However, compared to the visual system (15, 18), relatively

little is known about transcriptional changes occurring duringauditory plasticity. The contributions of cell type-specific tran-scriptional programs to network-level plasticity are poorly un-derstood in A1. For example, it remains unknown to what extentperineuronal nets (PNNs) enwrapping mature PV cells or myelinsignaling contribute to the closure of tonotopic plasticity as theydo for ocular dominance (19, 20). With the advancement ofsingle-cell RNA sequencing techniques, studies of cortical plas-ticity are further primed to uncover genetic programs with pre-viously unexplored depth across entire circuits without bias to

Significance

Early life acoustic experience shapes the organization andfunction of the primary auditory cortex (A1), but molecularmechanisms underlying these critical periods for auditoryplasticity are poorly understood. In this study, we use single-nucleus transcriptomics to characterize the multicellular geneexpression program in developing A1 and its regulation bytone exposure. We then functionally validated candidateplasticity triggers and brakes to reveal that strengthening ofinhibition initiates tonotopic plasticity, while the downstreammaturation of myelin-related signaling is associated with crit-ical period closure. These results both confirm conservedmechanisms and identify targets for the regulation of corticalplasticity.

Author contributions: B.T.K., E.E.D., M.E.G., and T.K.H. designed research; B.T.K., T.R.B.,E.E.D., and E.J.Z. performed research; B.T.K., T.R.B., E.E.D., and E.J.Z. analyzed data; andB.T.K., E.E.D., M.E.G., and T.K.H. wrote the paper.

Reviewers: D.F.C., Queen Mary University of London; and É.d.V.-S., McGill University.

The authors declare no competing interest.

This open access article is distributed under Creative Commons Attribution-NonCommercial-NoDerivatives License 4.0 (CC BY-NC-ND).

Data deposition: All sequencing data are available in the Gene Expression Omnibus (GEO)(accession no. GSE140883).1B.T.K., T.R.B., and E.E.D. contributed equally to this work.2Present address: Department of Biomedicine, Basel University, 4056 Basel, Switzerland.3To whom correspondence may be addressed. Email: michael_greenberg@hms.harvard.edu or hensch@mcb.harvard.edu.

This article contains supporting information online at https://www.pnas.org/lookup/suppl/doi:10.1073/pnas.1920433117/-/DCSupplemental.

First published May 13, 2020.

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https://orcid.org/0000-0001-8650-0986https://orcid.org/0000-0003-1380-2160http://crossmark.crossref.org/dialog/?doi=10.1073/pnas.1920433117&domain=pdfhttps://creativecommons.org/licenses/by-nc-nd/4.0/https://creativecommons.org/licenses/by-nc-nd/4.0/http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE140883mailto:michael_greenberg@hms.harvard.edumailto:michael_greenberg@hms.harvard.edumailto:hensch@mcb.harvard.eduhttps://www.pnas.org/lookup/suppl/doi:10.1073/pnas.1920433117/-/DCSupplementalhttps://www.pnas.org/lookup/suppl/doi:10.1073/pnas.1920433117/-/DCSupplementalhttps://www.pnas.org/cgi/doi/10.1073/pnas.1920433117

cell type (21, 22). Therefore, we utilized single-nucleus RNA-sequencing (snRNA-seq) in A1 across developmental timepoints spanning the tonotopic critical period in mice (postnataldays P10, P15, and P20) (9) with normal or altered acousticexperience in order to build a model for gene network changesbased on single-cell transcriptomics (23).

ResultsApplication of snRNA-Seq to the Developing Mouse Auditory Cortex.We performed snRNA-seq to generate a multicellular map ofgene expression in A1 at P10, P15, and P20 (Fig. 1A). A1 tissuewas freshly dissected and flash-frozen, bilateral hemisphereswere combined, and nuclei were isolated (Fig. 1B and Materialsand Methods) and then captured, and their mRNA was barcodedusing the inDrops platform (21). After initial quality filtering(>500 genes detected per nucleus), the dataset of developmentalsamples contained 31,293 nuclei, detecting on average 1,913transcripts (unique molecular identifiers [UMIs]) and 1,244 genesper nucleus (see SI Appendix, Fig. S1, for quality control metrics).Unsupervised clustering analysis identified 29 clusters with

distinct transcriptional profiles (22). We used canonical markergenes to classify nuclei into eight main cell types: excitatoryneurons (Slc17a7+), inhibitory neurons (Gad1+), oligodendro-cytes (Olig1+), astrocytes (Aqp4+), endothelial cells (Cldn5+),and microglia (Cx3cr1+) (Fig. 1C). We performed differentialgene expression analysis within cell types to explore develop-mental patterns of transcription across the tonotopic criticalperiod. Genes with a false discovery rate (FDR)

their expression pattern in tone-reared animals (Fig. 2E). Abun-dant gene expression changes exhibited by oligodendrocytes sug-gest they could be key mediators of transiently enhanced mapplasticity and delayed critical period closure (19, 20).One strength of single-nuclear sequencing is the ability to gain

novel insight into nonneuronal populations that may have beenpreviously underappreciated. This is particularly important giventhe growing recognition of the role of nonneuronal cells in circuitdevelopment, synapse refinement, and plasticity. However, the

role of astrocytes and microglia in the regulation of A1 criticalperiod remains unexplored. In V1, dark-rearing impairs thematuration of astrocytes (46), and the microglial P2Y12 purinergicreceptor is required for ocular dominance plasticity (47). Herewe found that glia exhibited a distinct transcriptional responseto tone-rearing during the tonotopic critical period. Comparedto controls, astrocytes from tone-reared animals demonstratedsignificantly higher expression of immediate-early genes, in-cluding Fos, Junb, and Nr4a1, as well as up-regulation of the

Fig. 1. Single nucleus sequencing of A1 across the tonotopic critical period. (A) Schematic of experimental design and time points for tissue collection. (B)Diagram of dissection approach used for auditory cortex tissue collection. (C) t-SNE plot of cells collected at P10, P15, and P20 under normally reared con-dition. Colors indicate different cell types. (D) Volcano plots depicting differentially expressed genes between P10 and P20 under normally reared conditionsin excitatory neurons, inhibitory neurons, and oligodendrocytes. Blue indicates statistically significant genes (FDR < 5%). (E) Gene ontology (GO) categoriesenriched in cell type-specific differentially expressed genes across A1 development. (F) t-SNE plots depicting c-fos and nr4a1 expression in P10, P15, and P20normally reared cells. There is an enrichment of c-fos and nr4a1 positive cells at P20. (G) Box plots of the trajectory of average gene expression for Gad1, Gad2,and Pvalb in inhibitory cells from P10 to P20 under conditions of normal development and tone rearing from P12 to P15. Box ranges represent the 25th and75th percentiles, while the box whiskers indicate the 95% confidence interval. Mean normalized gene expression is indicated. Pairwise gene expressionchange significance is indicated by asterisks (*FDR < 0.05, ***FDR < 0.001).

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Fig. 2. Cell type-specific effects of tone rearing during the tonotopic critical period. (A) t-SNE plots of cells from normally reared and tone reared mice at P15and P20. (B) Volcano plots depicting differentially expressed genes normally reared and tone reared conditions at P15 in excitatory neurons, inhibitoryneurons, and oligodendrocytes. Blue indicates statistically significant genes (FDR < 5%). (C) Gene ontology (GO) categories enriched in cell type-specificdifferentially expressed genes at P15 between normally reared and tone reared conditions. (D) Boxplot of the trajectory of average gene expression for Mbpfrom P10 to P20 under conditions of normal development and tone rearing from P12 to P15. Box ranges represent the 25th and 75th percentiles, while thebox whiskers indicate the 95% confidence interval. Mean normalized gene expression is indicated. Pairwise gene expression change significance is indicatedby asterisks (**FDR < 0.01, ***FDR < 0.001). (E) Boxplot of the trajectory of average gene expression for Mme and Bcan from P10 to P20 under conditions ofnormal development and tone rearing from P12 to P15. Box ranges represent the 25th and 75th percentiles, while the box whiskers indicate the 95%confidence interval. Mean normalized gene expression is indicated. Pairwise gene expression change significance is indicated by asterisks (*FDR < 0.05,***FDR < 0.001).

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Notch1 target Hes5. Notch signaling is thought to be importantfor neural activity-driven astrocyte maturation and morphologicalresponse to stimuli (48, 49).The excitatory amino acid transporter Slc1a3, as well as the

Kcnn2 small conductance calcium-activated channel, were bothdown-regulated in tone-reared astrocytes, perhaps as a result ofan activity-dependent compensatory mechanism. Analysis ofgene expression in microglia was limited by small cell numbers,but those from tone-reared mice demonstrated a nearly fourfoldreduction in complement gene C1qb and Fc-receptor–like mol-ecule Fcrls, both of which are typically down-regulated uponmicroglial activation (50, 51). Additional studies are needed todefine the functional significance of specific glial populations toauditory development and plasticity.

Functional Implications of Inhibitory Maturation. Extensive work inV1 has shown the functional maturation of GABAergic in-nervation is an important driver of critical period progression(52). In the absence of GAD65, a key GABA biosynthetic enzyme,a permanent precritical period state persists in V1 (53). Plasticitycan be rescued by administration of Diazepam, a benzodiazepineagonist which enhances particular GABAA receptor currents suchthat residual GABA levels in the absence of GAD65 drive suffi-cient inhibitory transmission. For example, while GAD65 knock-out mice do not exhibit a shift in ocular dominance following briefmonocular deprivation, these mice can exhibit plasticity at any agewhen treated with Diazepam (54).Our data showed that expression of Gad1 and Gad2, encoding

the two GABA synthetic enzymes GAD67 and GAD65, re-spectively, increased over development in A1 interneurons(Fig. 1G). One further theme that emerged from our A1 snRNA-seq data was the dynamic nature of inhibitory receptors acrossthe tonotopic critical period. This suggests that the maturation ofinhibitory tone may also be important for critical period timing inthe developing auditory cortex. We, therefore, examined howbroad manipulation of inhibitory transmission might affect A1topography and plasticity using voltage-sensitive dye imaging (9).Functional thalamocortical connectivity can be measured in an

acute slice preparation, where focal stimulation to single sites inauditory thalamus (ventral medial geniculate body [MGBv])elicits topographic responses in A1 (Fig. 3A). Our previous workrevealed that prior to the critical period at P10, focal electricalstimulation of medial MGBv sites (which receive high auditoryfrequency input in vivo) is more effective at driving A1 responsesthan lateral, low-auditory frequency sites (9). This bias is grad-ually lost over critical period development, with stimulation at allsites in MGBv eliciting similar maximal responses at topographiclocations across A1. In GAD65 knockout animals, this matura-tion failed to occur, and rostral sites continued to show greateractivation beyond the normal developmental window (Fig. 3B).Furthermore, in wild-type (WT) mice over early development,stimulation to single sites in MGBv typically yielded pro-gressively shorter latency responses, translocating from layer VIto layer IV (Fig. 3 C and D). In GAD65 knockout mice, the siteof shortest latency remained in deeper layer VI despite anoverall shortening of response latencies comparable to WTanimals (Fig. 3D).To test whether GAD65 is essential for the onset of the

tonotopic critical period, knockout mice were reared under re-peated 7-kHz tones between P12 and P15, which typically yieldsa shift in the tonotopic map and thalamocortical functionalconnectivity in control mice (Fig. 4 A and B) (9). The relation-ship between the stimulus site in MGBv and the site of maximalresponse in A1 is defined as the topographic slope and is equal to1 in control animals but drops in animals tone-reared during thecritical period (Fig. 4 B and C) (9). Mice lacking GAD65 failedto shift their topographic functional connectivity (Fig. 4 B and C)unless pretreated with Diazepam (Fig. 4D), consistent with the

hypothesis that their critical period onset is delayed (Fig. 3D).Tone exposure during a more extended time frame—from P8 toP20—also did not alter the tonotopic map in GAD65 null mice(Fig. 4C). In WT animals, administration of Diazepam givenprior to the natural critical period was also effective at drivingplasticity (Fig. 4E), shifting plasticity earlier and preventing itduring the expected window (Fig. 4F). These results collectivelydemonstrate that the development of inhibitory tone is necessaryfor the onset of critical period plasticity in A1, as it is in V1.

Molecular Brakes on Auditory Plasticity. In order to solidify changesmade to sensory maps during the critical period, molecularbrakes turn on to actively close the window and prevent furthercircuit refinements (55). Brakes are endogenous factors that haltor restrict plasticity, such as the maturation of the extracellularmatrix as a structural barrier to circuit rewiring. We found thatthe expression of PNN-related genes, including proteoglycansand proteases, were dynamically regulated across A1 develop-ment and in response to tone exposure (Fig. 2E). Notably,chondroitin sulfate proteoglycans (CSPGs) tightly enwrap PVbasket cells in the form of PNNs (19, 56). PNN intensity, asrevealed by Wisteria Floribunda Agglutinin (WFA) labeling, in-creased in A1 from P10 to P20 (Fig. 5 A and D, Left).Another example of a molecular brake is the increased mye-

lination of axons in cortex, which limits axon regrowth potentialand synaptic plasticity (57). Myelin-related gene expression andintracortical myelin basic protein (MBP) staining in WT miceincreased dramatically across this 10-d window (Fig. 5 A and C,Left). The GAD65 knockout animals displayed lower gray matterstaining intensity for both WFA and myelin (Fig. 5 C and D,Right), consistent with the shortest latency thalamocortical re-sponses remaining in layer VI (Fig. 3D). Diazepam treatment,which enabled normal tonotopic plasticity (Fig. 4F), did not fullyrescue PNN intensity by P20 (Fig. 5D) but returned myelin sig-nals to WT levels (Fig. 5C). Thus, myelination may serve as anearlier signal for map consolidation.Much of the myelin-related brakes on plasticity have been

attributed to signaling through the Nogo receptor, NgR (58).The neurite outgrowth inhibitor Nogo-A (Rtn4), which signalsthrough NgR, was up-regulated by oligodendrocytes at P20 (SIAppendix, Fig. S2C) but down-regulated in Tubb3+ neurons (59)after the critical period, which was prevented by tone-rearing(Fig. 5 E and F). This suggests a dynamic interplay betweenneuronal and glial populations to regulate both the level andtiming of plasticity within cortical circuits.Conveniently, the NgR is a key mediator of the downstream

response to several brake-like factors, including CSPGs, myelinfactors, and Nogo-A (Fig. 5G) (60). We thus examined whetherthe degree or timing of map plasticity in A1 might be altered byloss of this receptor. We found that NgR knockout animals wereplastic during both the normal critical period and beyond(Fig. 5H). Thus, myelin/CSPG-mediated signaling via the NgR isnecessary to restrict plasticity at the end of the tonotopic criticalperiod in A1. A permissive environment for structural changes inthe absence of NgR would allow a prolonged window for ana-tomical consolidation of functional refinements.

DiscussionEarly postnatal acoustic experience shapes the structural andfunctional organization of the auditory system (7). Neural ac-tivity during narrowly defined critical periods drives frequencyselectivity and tonotopic organization. Our study sought tocharacterize the cell type-specific transcriptional underpinningsof this carefully orchestrated multicellular collaboration. Weobserved gene expression changes in all major cell types acrossearly A1 development, during which these sensory-driven circuitrefinements occur. In addition to the vast transcriptomic re-source created in this study, we also extend findings from the

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visual system to the auditory cortex. Specifically, we demonstratethat GAD65, a key synthetic enzyme for GABA, and the Nogoreceptor, a mediator of myelin/CSPG signaling, are both neces-sary for proper critical period onset and offset, respectively (20,

53). These results underscore conserved mechanisms for theregulation of cortical plasticity.Our study further reveals an activity-dependent mobilization

of molecular machinery enabling plasticity. Thus, tone-rearing

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Fig. 5. Critical period closure in A1 signaled by myelin/PNN formation and NgR. (A) Schedule for staining (arrows) with reference to typical A1 critical periodin WT mice. (B) Schedule for staining (arrows) with reference to Diazepam or vehicle treatment in GAD65−/− mice. (C and D) (Left) Quantification of relativeMBP (C) and WFA (D) staining intensity in P10 (n = 5, 4), P13 (n = 5, 4), P16 (n = 4, 4), and P20 (n = 4, 4) WT mice. **P < 0.01 ***P < 0.001, two-way ANOVAwith post hoc Bonferroni correction; mean ± SEM. (Right) Quantification of MBP (C) and PNN (D) staining in P20 WT nonexposed (black, n = 4, 4), GAD65−/−

(red, n = 5, 3) and GAD65−/− injected with DZ between P12-P15 (blue dashed, n = 5, 4). *P < 0.05; **P < 0.01, two-way ANOVA with post hoc Bonferronicorrection; mean ± SEM. (E) Fluorescent in situ hybridization (FISH) in A1 labeling cellular nuclei (DAPI), Nogo-A (Rtn4), neuronal marker (Tubb3), and theirmerged images across ages and tone-rearing. Representative images are from Layer V/VI. Red outlines are QuPath (67) segmented nuclei and an expandedestimated cell border. (Scale bar, 10 μm.) (F) Quantification of mean FISH signal intensity in Tubb3+ nuclei (Materials and Methods) at P10, P20, and P20 aftertone-rearing during the critical period (P12 to P15). Quantification was averaged across all layers. ***P < 0.001, Kruskal–Wallis rank sum test with post hocDunn test; horizontal lines in violin plot indicate quantiles 0.25, 0.50, and 0.75. (G) Schematic of putative NgR ligands in myelin membrane (Nogo-66, OMgp,and MAG) and the extracellular matrix (CSPG). (H) Schedule for sound exposure and VSDI recordings (arrows). Topographic slopes for NgR−/− mice exposedto 7 kHz between P8 and P11 (P11, white, n = 5), between P12 and P15 (P15, white, n = 8) or between P16 and P19 (P19, red, n = 4). **P < 0.01, t test;mean ± SEM.

11750 | www.pnas.org/cgi/doi/10.1073/pnas.1920433117 Kalish et al.

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per se altered normal developmental trajectories of bothGABAergic circuits and structural barriers to plasticity such asmyelination and PNNs. The idea that abnormal acoustic expe-rience during the critical period alters maturation trajectorieshas been suggested by other studies. For example, GABAB-mediated inhibitory long-term depression (iLTD) is triggered byprepost pairing of action potentials at PV-to-principal cell syn-apses during the A1 critical period, which is thought to underlie adisinhibitory mechanism permissive for plasticity (61). This iLTDswitches to potentiation (iLTP) as development proceeds.After tone-rearing, the number of cells responsive to the

rearing frequency increases in a topographic zone of A1, andthey exhibit a premature switch to iLTP (61). Instead, our datafrom A1 biopsies show that the gene encoding GABAB subunit 2normally decreases over early development but increases in tone-reared animals, which should mediate continued iLTD. Onepossibility is that tone-rearing drives both iLTP and iLTD in atopographic manner to mediate map expansion to the rearingfrequency and compensatory retraction in others. In addition,presynaptic GABAB receptors have been shown to regulate theexperience-dependent switch from depression to facilitation ininhibitory plasticity (62).The auditory system is a tractable model of experience-

dependent plasticity due, in part, to its topographic organiza-tion. The use of snRNA-seq allows for the dissection of layer-specific excitatory cell types and classes of inhibitory neurons, aswell as nonneuronal cells. However, this approach homogenizesthe tonotopic organization of cells in A1. It is not clear whetherdistortion of inhibitory maturation or molecular brake onsetoccurs within the tone-responsive areas of cortex in response tooverstimulation or if the neighboring part of the tonotopic map issilenced in a competitive manner. Future studies could usemultiplexed fluorescence in situ hybridization to visualize spa-tially restricted cell type-specific transcriptional changes toaddress these questions.Our data represent a significant advance over existing re-

sources as this study profiles transcription with cell type speci-ficity across time and with critical period perturbation. However,the use of snRNA-seq also has several limitations. First, thisapproach has low capture efficiency, such that a small proportionof a cell’s total transcriptome is represented in the final se-quenced library. This challenge makes it difficult to distinguishbetween biologic variability and technical noise for low-abundance transcripts, such as the nicotinic acetylcholine brakeLynx1 (25).The low amount of input material also leads to high levels of

technical noise, again making it difficult to observe biologicvariation. Owing to low capture efficiency and stochastic geneexpression, gene dropout (where gene expression is zero in agiven cell) is quite common, leading to zero-inflated expres-sion data. Other potential sources of bias include the tissue

dissociation method, as enzymatic treatments may affect cellviability, as well as the low number of animals from which nucleiwere collected. The relative merits of single-cell sequencing, asopposed to bulk RNA sequencing, depend on many factors, in-cluding the specific scientific question, cell type abundance, andgene-of-interest expression level.Despite these limitations, snRNA-seq is an important discov-

ery tool with which to obtain previously impossible degrees ofcellular resolution. For example, we found that Nrgn is up-regulated in tone-reared interneurons (SI Appendix, Fig. S2B),while this gene is restricted to principal cells in the mature cortexacross species (63). There is precedent that Nrgn can be tran-siently expressed in GABAergic interneurons in a developmen-tally restricted fashion (64) and in subsets of GABAergicneurons in other contexts (65). Similarly, Kv3.1 may be tran-siently expressed in oligodendrocyte precursor cells at evenearlier ages (66). This highlights the importance of unbiasedexamination of longitudinal trajectories of dynamic gene ex-pression across cell types. Overall, our results are consistent witha pivotal role for PV+ circuits in regulating critical period pro-files across brain regions (32). The transcriptomic data obtainedin the present study provide insights into critical period regula-tion and a resource for future investigation into cell type-specificregulatory mechanisms in auditory cortex development.

Materials and MethodsAll experiments using animals were performed according to protocols ap-proved by the Harvard University Institutional Animal Care and Use Com-mittee. Tone rearing was performed between postnatal days 12 to 15, andlitters were moved back to standard housing on postnatal day 16. The au-ditory cortex was dissected and flash frozen; a nuclear suspension wassubsequently prepared using gradient centrifugation. Single nuclei werecaptured, barcoded, and sequenced according to the inDrops technique aspreviously described (21). All sequencing data are available in the GeneExpression Omnibus (GSE140883). Voltage-sensitive dye imaging of C57BL/6J, GAD65−/−, or NgR−/− mice was performed on acutely prepared auditorythalamocortical slices, as described previously (5, 9). Additional details arecontained in SI Appendix, Extended Experimental Procedures.

ACKNOWLEDGMENTS. We thank members of the M.E.G. and T.K.H.laboratories for discussions and critical reading of the manuscript. We thankMari Nakamura, MSc, for expert mouse colony maintenance and MarykateO’Malley, MSc, BSN, RN, for her assistance with figures. We thank the Neu-robiology Department and Neurobiology Imaging Facility for consultationand instrument availability supported in part by the Neural Imaging Centeras part of an National Institute of Neurological Disorders and Stroke P30Core Center grant NS072030. We further thank the Single Cell Core at Har-vard Medical School, Boston, MA, for performing the single-cell RNA-Seqsample preparation. This work was funded by NIH grant R01 NS028829 toM.E.G., a Pediatric Scientist Development Award and March of Dimes fund-ing to B.T.K., the Harvard Society of Fellows to T.R.B., Thomas T. HoopesPrize to E.J.Z., and a National Institute of Mental Health Silvio Conte Center(P50MH094271) andWorld Premier International Research Center for Neuro-intelligence (Japan Society for the Promotion of Science) grant to T.K.H.

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